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Department of Animal and Poultry Science, University of Guelph, Ontario, Canada N1G 2W1
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Abstract |
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 Top Abstract INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONLITERATURE CITED |
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Key Words: conjugated linoleic acid • odd- and branched-chain fatty acid • rumen bacteria and protozoa • vaccenic acid
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INTRODUCTION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONLITERATURE CITED |
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). In the rumen, dietary lipids are subject to hydrolysis by microbial lipases followed by biohydrogenation of the unsaturated FFA by rumen bacteria. In the case of linoleic acid (18:2n-6), the end product of the hydrogenation is stearic acid (18:0; Jenkins, 1993). When biohydrogenation does not go to completion, intermediates (e.g., trans-18:1) from the incomplete biohydrogenation of PUFA become available for deposition in microbial biomass as well as in animal tissues (Bauman and Griinari, 2003).
One of the microbial transformations in the rumen is the microbial synthesis of odd- and branched-chain fatty acids (Kim et al., 2005). These fatty acids are regarded as important compounds within microbial lipids (O’Kelly and Spiers, 1991) to maintain optimal fluidity of the microbial cell membrane. Branched-chain fatty acids of bacterial origin can make up 1.0 to 3.0% of milk lipids (Alonso et al., 1999) and carcass lipids (Bas and Morand-Fehr, 2000).
On the other hand, there has been a great deal of interest in possible health benefits of cis-9, trans-11 CLA in human diets (Ip et al., 1999). The major sources of CLA for humans are ruminant products such as meat, milk, and other dairy products (Chin et al., 1992). The major CLA, cis-9, trans-11 CLA, is synthesized either in the rumen as an intermediate in the biohydrogenation of linoleic acid or in the tissues by 
-desaturase from vaccenic acid (VA; 18:1 trans-11), another intermediate in ruminal biohydrogenation (Griinari and Bauman, 1999). Formation of CLA in the rumen has been mainly associated with bacterial activity, and the role of protozoa in the synthesis of CLA is unknown. Rumen protozoa contain a greater proportion of unsaturated fatty acids than bacteria (Devillard et al., 2004). Approximately 7.5 to 15% of the lipids present within the rumen digesta are of protozoal origin (Keeney, 1970). Thus, protozoa represent a potential source of lipid for the host animal (Katz and Keeney, 1967).
Knowledge of the fatty acid profile of microbial lipids is of great nutritional importance to animals and, subsequently, their products. The purpose of this research was to determine the fatty acid composition of mixed ruminal bacteria and protozoa with emphasis on CLA, VA, and odd- and branched-chain fatty acids.
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MATERIALS AND METHODS |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONLITERATURE CITED |
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Fractionation of Rumen Contents
Three, dry, ruminally fistulated Holstein cows (719 ± 39 kg of BW) were given ad libitum access to a total mixed ration (Table 1) offered twice daily at 0800 and 1500 h for 3 wk before the first rumen sampling. Ad libitum intake was insured by feeding up to 5% orts. The mean intake was 17.5 ± 1.8 kg of DM/d. Fresh H2O was available at all times. Rumen contents were collected from the 3 cows 3 times at 3-d intervals before the morning feeding. The samples were composited during each collection, transported to the laboratory, blended in a kitchen blender (Kenmore mini food processor model CH-03C, Sears Canada, Toronto, Ontario, Canada) for 20 s to detach the microbes from the feed particles, and then strained through 2 layers of surgical gauze into a 2-L separatory funnel flushed continuously with CO2. Strained rumen contents (1 L) were then diluted with a 30% volume of MB9 buffer (2.8 g of NaCl/L; 0.1 g of CaCl2·2H2O/L; 0.1 g of MgSO4·7H2O/L; 2.0 g of KH2PO4/L; and 6.0 g of Na2HPO4/L; Or-Rashid et al., 2001). The funnel containing rumen fluid was left to stand for 60 to 90 min in a H2O bath (37°C) to allow small feed particles to float and the microbial fraction to sediment to the bottom of the funnel.
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and Lee et al. (2000). Briefly, most of the lower liquid portion (not the floating feed particles) was collected gravimetrically from the funnel and centrifuged at 150 x g for 5 min at 4°C, and the supernatant was designated ruminal bacteria plus a few protozoa. A portion of this supernatant was further centrifuged at 500 x g for 5 min at 4°C, and the resultant supernatant (ruminal bacterial preparation) was found by microscopic examination to be free from all protozoa and virtually all plant material. Some of this preparation was then centrifuged at 12,000 x g for 30 min at 4°C, and the resultant residue was resuspended in MB9 buffer and labeled bacterial suspension. The pellet obtained from the initial 150 x g centrifugation was used to produce the rumen protozoal preparations. The pellet was diluted with MB9 buffer and washed by centrifugation (approximately 4 to 6 times) until essentially free from feed material (checked under microscope) and finally resuspended in the same buffer to give a protozoal suspension. Bacterial and protozoal pellets resuspended in MB9 buffer were then transferred into 2-mL Eppendorf tubes and kept at –20°C until further analysis.
Preparation and Derivatization of the Samples for Analysis of Fatty Acids
The suspended microbial pellets in the Eppendorf tubes were thawed, 1 g of zirconium beads [0.5 and 0.1 mm at a ratio of 1:1 (wt/wt)] was added, and the tubes were shaken for 2.5 min twice, with a 5-min intervening period, at room temperature on a reciprocating shaker (Mini-Bead-Beater-8, Biospec Products Inc., Bartlesville, OK) at the maximum speed to cause mechanical disruption of the ruminal contents. Contents of the tubes were then transferred into 15-mL culture tubes equipped with screw caps and Teflon liners. Two portions of 1.9 mL of CH3OH were added to the Eppendorf tube to ensure complete transfer. The contents of the culture tubes were well mixed by vortexing and allowed to stand at room temperature for 1 h. After 1 h, 1.9 mL of CHCl3, 1.8 mL of H2O, and 0.1 mL of 3 M HCl were added, vortex-mixed, and centrifuged. The acid was added to ensure the pH of the extract was acidic. The CHCl3 layer (bottom phase), containing esterified lipid and FFA, was removed using 2 pasture pipettes, one inserted into another. The CH3OH-H2O phase was extracted with an additional 1.9 mL of CHCl3, and the CHCl3 phases were combined, dried over anhydrous Na2SO4, and filtered. Chloroform was removed from the vials under a stream of N2, and 3 drops of C6H6 was added and mixed. Then, 1 mL of 1 M NaOH (dissolved in 95% C2H5OH) was added to the vials, vortex-mixed, and kept at room temperature overnight in a dark place.
The following day, 1.0 mL of 3 M HCl was added and extracted twice with C6H14 (1.0 mL each time). After evaporation of C6H14 under N2, 0.8 mL of C6H6, and 0.2 mL of CH3OH were added. After mixing, 7 to 8 drops of trimethylsilyl-diazomethane were added to the vials, shaken lightly (trimethylsilyl-diazomethane is explosive) and occasionally over 60 min. After derivatizing for 60 min at room temperature, a few drops of glacial acetic acid were added until the yellow color disappeared to inactivate the derivatizing agent. Then, 0.5 mL of H2O and 1.0 mL of C6H14 were added and vortexed. The C6H14 layer was extracted, and this was repeated 1 time with same amount of C6H14. The vial containing C6H14 (approximately 2 mL) was placed under a stream of N2 until 90 to 100 µL remained. Then, the C6H14, containing fatty acid methyl esters, was analyzed by GLC. For mechanical disruption of the microbial cells, we did not use a sonicator as commonly used for disrupting cells because Or-Rashid et al. (2003) suggested that it destroyed 50 to 60% of the total unsaturated fatty acids.
Analysis of Fatty Acid Composition by GLC
Analysis of fatty acid methyl esters was performed using an Agilent 6890N GLC (Agilent Technologies, Palo Alto, CA) equipped with a split-splitless injector at 250°C, a flame ionization detector at 250°C, and a CP Sil 88 column (100 m x 0.25 mm, 0.2 µm of film thickness, Varian Inc., Mississauga, Ontario, Canada). Hydrogen was used as the carrier gas at a constant flow rate of 1 mL/min. The temperature of the GLC oven was set to 45°C for 4 min, increased at 13°C/min to 175°C and held for 27 min, and increased at the rate of 4°C/min to a final temperature of 215°C and held for 35 min. Agilent Technologies Chemstation software (Rev. B.01.01) was used for data analysis.
A 1-µL sample was injected at splitless mode. Peaks were routinely identified by comparison of the retention times with fatty acid methyl ester standards (GLC no. 463, no. UC-59-M, 21:0, 23:0, and 26:0, Nu-Check-Prep Inc., Elysian, MN). Individual isomers of 18:1 fatty acids were determined as follows. The temperature of the GLC oven was maintained at 45°C for 4 min, increased to 167°C at a rate of 13°C/min and held for 40 min, and again increased at the rate of 4°C/min to a final temperature of 218°C and held for 23 min. Peaks of 18:1 isomers were identified by order of elution according to Precht and Molkentin (1997), Shingfield et al. (2003), and Loor et al. (2004). Fatty acid composition was expressed as a percentage of total fatty acids.
Chemical Analyses
Feed and orts samples were collected 3 times per week and analyzed for DM content by drying them in an oven at 60°C for 48 h (AOAC, 1990). A subsample was ground using a Wiley mill with a 1-mm screen (Thomas-Wiley, Philadelphia, PA) and stored at –20°C until analyzed. The feed samples were analyzed for CP using the Kjeldahl procedure (AOAC, 1990) and for ADF (AOAC, 1990), NDF (Goering and Van Soest, 1970), ash (AOAC, 1990), and crude fat (AOAC, 1990).
Statistical Analysis
The data were analyzed as a completely randomized design using PROC MIXED (SAS Inst. Inc., Cary, NC) with the model Yij = µ + 
i + ßj + 
ij, where µ = the overall mean; = the effect of day (i = 1 to 3); ß = the effect of treatment (j = bacteria or protozoa); and ij = the residual error. Effects were considered significant at a probability of P < 0.05. Means of individual fatty acids were the average of 3 measurements.
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RESULTS AND DISCUSSION |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONLITERATURE CITED |
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). The results of fatty acid analyses are summarized in Tables 2, 3, and 4. The predominant fatty acids were palmitic (16:0) and stearic (18:0) in both the bacterial and protozoal fractions. Palmitic acid was 74% greater in the protozoal than in bacterial fatty acids (P < 0.001), whereas bacteria had 2.25-times greater stearic acid (18:0) content compared with protozoa (P = 0.009). These findings are in agreement with earlier work (Williams and Dinusson, 1973). Emmanuel (1974) showed that mixed protozoal suspensions formed structurally related long-chain fatty acids, principally 16:0 from short-chain precursors such as acetate and butyrate. Protozoa can also incorporate 16:0 from feed supplied to the host animals (Williams, 1986). In contrast, Wu and Palmquist (1991) showed that rumen bacteria more likely incorporated palmitic acid from feed, sparing de novo synthesis from short-chain precursors (e.g., acetate and butyrate), and this might explain the greater percentage of 16:0 in protozoa compared with bacteria.
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). These fatty acids are synthesized by chain elongation of branched chain VFA (i.e., branched short-chain carboxylic acids) generated by metabolism of branched-chain AA (Garton, 1977). The branched-chain fatty acids could be considered as H acceptors during anaerobic fermentation, thus the synthesis of these long branched-chain fatty acids from the branched short-chain carboxylic acids (e.g., isobutyric, isovaleric, and 2-methylbutyric acids) may contribute to energy conservation (Bas et al., 2003).
On the other hand, 18:1 trans-11 was found in much greater (P = 0.001) amounts (approximately 3 times) in the protozoal fraction than in the bacterial fraction (6.6 vs. 2.0% of total fatty acids; Table 3). We hypothesize that rumen protozoa may have 11-desaturase activity, allowing conversion of 18:0 to 18:1 trans-11. In support of our hypothesis, it was shown by Emmanuel (1974) in a study of mixed rumen protozoa that octadecenoic acids, consisting mainly of the 11 isomer, were synthesized by direct desaturation of saturated acids. However, engulfment of VA (associated with feed particles and bacterial cells) by protozoa may be another way it is incorporated into protozoal lipids. The percentages of other trans-18:1 isomers such as trans-9, trans-10, and trans-12 were greater (P 
0.05) in protozoa than bacteria (Table 3).
The percentages of odd-chain fatty acids (9:0, 11:0, 17:0, 19:0, etc.) in bacteria were greater (P 0.05) than in protozoa, except for 13:0 (P = 0.47) and 15:0 (P = 0.12; Table 2). Microbial odd-chain fatty acids are formed through elongation of propionate or valerate (Emmanuel, 1974; Kaneda, 1991). Odd- or branched-chain fatty acids with an iso- or anteiso-structure provide biological advantage to rumen microbes because the odd-chain fatty acids and the iso-fatty acids with n atoms of C have a melting point 1 to 2°C lower than those of straight-chain fatty acids with n-1 atoms of C (Wu and Palmquist, 1991; Gunstone et al., 1994). The melting point of even-chain fatty acids with an anteiso-structure is lower by 25 to 30°C compared with its corresponding straight-chain SFA (Gunstone et al., 1994). In rumen microbes with a lower content of unsaturated fatty acids, these odd- and branched-chain fatty acids provide structural lipids with optimal fluidity for cell membranes (Hauser et al., 1979). However, the total amount of odd- and branched-chain fatty acids in the protozoal fraction was 47% lower than that of the bacterial fraction (Table 2). In this case, the 2- to 3-times greater content of unsaturated fatty acids in protozoa might compensate for the deficiency of available odd- and branched-chain fatty acids for maintaining lipid fluidity and cell membrane function of protozoa. Odd- and branched-chain fatty acids are generally rare or absent from feeds (Diedrich and Henschel, 1990). In fact, Keeney et al. (1962) suggested that major sources of these fatty acids in rumen and milk are from bacterial origin and might be used as markers to quantify bacterial matter leaving the rumen. Dewhurst et al. (2000) also suggested that these fatty acids in duodenal digesta and milk could provide a qualitative description of the proportions of different classes of microbes leaving the rumen. In this group of fatty acids, the protozoal fraction had 2.05-times greater anteiso-17:0 (2.89% of total fatty acids) content than bacterial fraction. Therefore, anteiso-17:0 content in milk and meat might be used as a marker to quantify protozoal biomass in response to dietary shifts. However, further research is necessary to correlate anteiso-17:0 content in milk or meat with changes of protozoal biomass with different diet formulations. Similarly, iso-13:0, iso-15:0, anteiso-15:0, etc. could be used to quantify bacterial biomass (Vlaeminck et al., 2006).
Among MUFA, oleic acid (18:1 cis-9) was the greatest both in bacteria and protozoa. Protozoa contained 3.7-times greater oleic acid than bacteria (P = 0.006; Table 3). The percentages of linoleic acid (18:2 cis-9, cis-12) were the greatest among PUFA both in bacteria and protozoa, followed by linolenic acid (18:3n3; Table 2). The protozoal fraction had 4.8-times greater linoleic acid (7.2% of total fatty acids) compared with the bacterial fraction (1.5%; P < 0.001). One of the causes is that the linoleic acid uptake rate from feed particles by protozoa might be greater than that by bacteria. However, Viviani and Borgatti (as cited by Viviani, 1970) demonstrated that the rumen protozoa were able to incorporate 14C-acetate into linoleic acid. Therefore, the linoleic acid synthesizing ability of protozoa could be another factor for increased level of this fatty acid in protozoal biomass. Additionally, bacteria contained more nervonic acid (24:1 cis-15) than protozoa (P = 0.04), although the percentage was very low in both fractions (Table 2).
The total SFA in bacteria were 29% greater than that in protozoa (P < 0.001; Table 2). However, the long-chain SFA (17:0 to 26:0) in bacteria were 123% greater than in protozoa (P = 0.004). Conversely, protozoa had greater percentages of total MUFA, total n-6 PUFA, total n-3 PUFA, and total CLA (2.3-, 3.9-, 3.45-, and 6.8-times, respectively) compared with bacteria. Thus, fatty acids in mixed rumen protozoa were less saturated than those of mixed rumen bacteria. In addition, bacteria had lower levels of total trans-18:1 isomers than did protozoa (3.6 vs. 9.7% of total fatty acids; P = 0.007). Protozoa had a greater proportion of total cis-18:1 than bacteria (8.9 vs. 3.2% of total fatty acids; P = 0.008).
Among all CLA isomers, cis-9, trans-11 CLA was the most abundant isomer in both bacteria and protozoa (Table 4). Interestingly, cis-9, trans-11 CLA proportions in protozoa were 8.6-times greater than bacteria (P = 0.002). It is not clear why protozoal lipid contained so much more cis-9, trans-11 CLA (1.32% of total fatty acids) than that of bacteria. However, protozoa may have 9-desaturase activity that could convert VA to cis-9, trans-11 CLA. Additionally, protozoa could incorporate cis-9, trans-11 CLA from symbiotic bacteria, which isomerize 18:2 cis-9, cis-12 to cis-9, trans-11 CLA inside the protozoal cells. At present, there are no reports demonstrating the capability of protozoa to isomerize 18:2 cis-9, cis-12 to cis-9, trans-11 CLA. However, earlier results, without showing isomerization, suggested that protozoa had limited or no ability to hydrogenate various substrates (Chalupa and Kutches, 1968; Girard and Hawke, 1978). The second largest CLA was the mixture of trans-9, trans-11 CLA plus trans-10, trans-12 CLA, which was present in both bacterial and protozoal fractions. Again, protozoa had greater percentages than bacteria (P = 0.01). Other minor CLA isomers detected in bacteria and protozoa included the following: cis-10, trans-12; trans-9, cis-11; trans-10, cis-12; and trans-11, trans-13.
It should be noted that our microbial samples were all collected before feeding. Fatty acid profiles of rumen microbial fractions (e.g., bacterial and protozoal biomass) could change during the postprandial period (Devillard et al., 2004). Diet variation is another factor that could affect the fatty acid profiles of rumen microbes, and the relevance of our observations might be limited to diets similar to those we fed.
In summary, the main fatty acid in mixed rumen bacteria was stearic acid, whereas palmitic acid was predominant within protozoa. Both bacteria and protozoa contained odd- and branch-chained fatty acids in different proportions. These fatty acids could play a role in H transfer, thus conserving potential energy. Protozoa had 2.05-times greater anteiso-17:0 content than bacteria; thus, this fatty acid may be a potential biomarker for protozoal biomass. Additionally, protozoa contained a greater proportion of unsaturated fatty acids and CLA than the bacteria. These results suggest that the presence of protozoa in the rumen may increase the supply of CLA and other unsaturated fatty acids for lower gut absorption by ruminants.
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Footnotes |
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2 Corresponding author: bmcbride{at}uoguelph.ca
Received for publication June 14, 2006. Accepted for publication November 21, 2006.
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LITERATURE CITED |
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TopAbstractINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONLITERATURE CITED |
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